regulation of growth anisotropy in well-watered and water

Regulation of Growth Anisotropy in Well-Watered andWater-Stressed Maize Roots. II. Role of Cortical Microtubules

and Cellulose Microfibrils1

Tobias I. Baskin*, Herman T.H.M. Meekes, Benjamin M. Liang, and Robert E. Sharp

Division of Biological Sciences (T.I.B., H.T.H.M.M.) and Department of Agronomy, Plant Science Unit(B.M.L., R.E.S.), University of Missouri, Columbia, Missouri, 65211

We tested the hypothesis that the degree of anisotropic expan-sion of plant tissues is controlled by the degree of alignment ofcortical microtubules or cellulose microfibrils. Previously, for theprimary root of maize (Zea mays L.), we quantified spatial profilesof expansion rate in length, radius, and circumference and thedegree of growth anisotropy separately for the stele and cortex, asroots became thinner with time from germination or in response tolow water potential (B.M. Liang, A.M. Dennings, R.E. Sharp, T.I.Baskin [1997] Plant Physiol 115:101–111). Here, for the same ma-terial, we quantified microtubule alignment with indirect immuno-fluorescence microscopy and microfibril alignment throughout thecell wall with polarized-light microscopy and from the innermostcell wall layer with electron microscopy. Throughout much of thegrowth zone, mean orientations of microtubules and microfibrilswere transverse, consistent with their parallel alignment specifyingthe direction of maximal expansion rate (i.e. elongation). However,where microtubule alignment became helical, microfibrils oftenmade helices of opposite handedness, showing that parallelismbetween these elements was not required for helical orientations.Finally, contrary to the hypothesis, the degree of growth anisotropywas not correlated with the degree of alignment of either microtu-bules or microfibrils. The mechanisms plants use to specify radialand tangential expansion rates remain uncharacterized.

During animal morphogenesis, cells grow, move, con-tract, or die, whereas in plant morphogenesis, cells onlygrow. A growing plant organ, to produce any other formthan a sphere, must grow at different rates in differentlocations or directions. When growth rates are differentand in different directions, growth is said to be anisotropic.Anisotropy is a nearly ubiquitous feature of plant growth,not only for the leaves of grasses and cylindrical organs

such as coleoptiles, stems, and roots, which expand prin-cipally in length, but also for laminar organs such as dicotleaves and petals, which may expand isotropically in theplane of the lamina but which expand minimally perpen-dicularly to the lamina. Growth anisotropy has even beenreported for tip-growing cells in the rare cases in whichboth axial and tangential growth components have beenresolved (Castle, 1958; Green, 1965).

Plants control expansion by controlling how the cell wallyields to turgor pressure. Because turgor pressure is iso-tropic, the cell wall can yield anisotropically only when themechanical properties of the cell wall are anisotropic. Themost prominent anisotropic component of the cell wall iscellulose. This polymer is synthesized at the plasma mem-brane as long chains of b-1,4-linked glucose residues thatassociate laterally into microfibrils (Brown et al., 1996).Microfibrils are usually aligned in parallel, which rein-forces the cell wall anisotropically and may guide thesubsequent assembly of polymers around the celluloseframework (Cosgrove, 1997). In the diffusely growing cellsof higher plants, the aligned deposition of microfibrils isthought to be dictated by the alignment of cortical micro-tubules (Cyr, 1994; Wymer and Lloyd, 1996).

Anisotropic expansion is characterized by two parame-ters: the direction in which the maximal expansion rateoccurs and the degree to which the maximal expansion rateexceeds the minimal rate. The direction of maximal expan-sion is specified by the direction of the cellulose microfi-brils, according to several lines of evidence. First, the axisof maximal expansion rate is usually perpendicular to thenet alignment of microfibrils (Green, 1980; Taiz, 1984).Second, growth anisotropy is reduced or even eliminatedwhen microfibril synthesis is inhibited chemically or ge-netically (Hogetsu et al., 1974; Arioli et al., 1998). Third,expansion becomes essentially isotropic when, during de-velopment or in response to inhibitors or hormones, mi-crofibrils are deposited without a predominant alignment(Richmond, 1983; Hogetsu, 1989; Iwata and Hogetsu, 1989;Sakaguchi et al., 1990). Finally, cell cultures can be obtainedthat have scant cellulose in their cell walls and that un-dergo turgor-driven enlargement (Shedletzky et al., 1990)but, to our knowledge, these never expand anisotropically.

In contrast to the direction of maximal expansion rate,there is almost no information about what controls thedegree of anisotropy. One reason for this lack is that radialor tangential expansion rates are rarely quantified and so

1 This paper is dedicated to the memory of Paul B. Green(1931–1998). This project was funded in part by grant no.94ER20146 (to T.I.B.) from the U.S. Department of Energy and doesnot constitute endorsement by that department of views expressedherein by the University of Missouri Research Board (award no.RB-95038 to T.I.B.), by the Cooperative States Research Service,U.S. Department of Agriculture (award no. 95-37100-1601 (withR.E.S. and W.G. Spollen), and by the University of Missouri Foodfor the 21st Century Program (R.E.S). This is a contribution fromthe Missouri Agricultural Experiment Station, journal series no.12,841.

* Corresponding author; e-mail [email protected]; fax 1–573– 882– 0123.

Plant Physiology, February 1999, Vol. 119, pp. 681–692, www.plantphysiol.org © 1999 American Society of Plant Physiologists

681

the degree of growth anisotropy is usually unknown. Anexception is the giant internode of characean algae, forwhich the ratio of growth rate in length to diameter isabout 4.5 (Green, 1964). However, this ratio remains con-stant during development, so that the studies of the role ofcellulose microfibrils in the control of expansion usingthese cells have not provided insight about variation in thedegree of growth anisotropy (Taiz, 1984).

There is a further difficulty that has limited our under-standing of how the degree of growth anisotropy is con-trolled in the organs of higher plants. For a single cell suchas an algal internode, growth anisotropy is characterizedby the changes in cell length and diameter; however, for acylindrical, multicellular organ such as a root, measure-ments of elongation and organ diameter over time are notsufficient. Although all tissues at a given distance from theapex must elongate at the same rate (otherwise the rootwould tear or cells would slip), different tissues, e.g. thestele and cortex, may expand in diameter at different rates.Moreover, expansion in diameter has two components,radial and tangential (i.e. circumferential), and these do nothave to be equal at a given location (Liang et al., 1997).Therefore, the degree of growth anisotropy may differbetween different tissues as well as between cell walls intangential and radial planes.

To our knowledge, the only study to have measuredrates of expansion in diameter separately for different tis-sues and to obtain both radial and tangential terms is thefirst paper in this series, concerning the control of growthanisotropy in the primary root of maize (Zea mays) (Lianget al., 1997). These roots become thinner with time fromgermination, reaching steady-state growth after about 3 d.We found for both the cortex and stele that neither radialnor tangential expansion rate was proportional to elonga-tion rate and, hence, unlike the giant algal internodes, thedegree of growth anisotropy varied. In fact, the degree ofanisotropy, calculated as the ratio of longitudinal to radial(or to tangential) expansion rate varied with time andbetween positions by more than 1 order of magnitude.Additionally, we analyzed directional expansion rates inroots exposed to a water-stress treatment, which had beenpreviously shown to cause the roots to thin (Sharp et al.,1988). The reduced diameter contributes significantly to themaintenance of root elongation under water stress becauseroot volume decreases as the square of the radius andhence less water and solutes are needed for growth (Sharpet al., 1990). The degree of growth anisotropy in water-stressed roots differed markedly from well-watered roots,being increased in the apical region of the growth zone anddecreased in the basal region.

As an approach to understanding what determines thespatial profiles of expansion rate and anisotropy, the ob-jective of this study was to quantify the alignments ofmicrotubules and microfibrils as a function of position inboth well-watered and water-stressed roots. Our resultsbear on three related questions. First, what is the relation-ship between the alignments of microtubules and microfi-brils? These alignments have been compared most often inthe epidermis of stems; however, in this tissue comparisonsare hindered by the orientations continually shifting

among transverse, longitudinal, and oblique. Second, iselongation rate controlled by the alignments of microtu-bules or microfibrils? Because both are generally observedto be transverse during rapid elongation and oblique (orlongitudinal) otherwise, the alignment of these elementshas been suggested to control elongation rate. Third, whatcontrols the degree of growth anisotropy? The answer hasbeen hypothesized to be the degree of alignment amongcellulose microfibrils, but this hypothesis has not beentested decisively. Answering this question is essential be-cause until we know how cells expand anisotropically, wewill not understand plant morphogenesis.

MATERIALS AND METHODS

Seeds of maize (Zea mays L. cv FR27 3 FRMo17) weregerminated, then transplanted into vermiculite at a waterpotential of approximately 20.03 MPa (well-watered) orapproximately 21.63 6 0.08 MPa (water-stressed, mean 6sd, n 5 20; water potential was measured in every exper-iment), and grown in darkness at 29°C and near-saturationhumidity, as described by Liang et al. (1997). Well-wateredroots were harvested at 24 or 48 h and water-stressed rootswere harvested 48 h after transplanting. Primary roots usedfor all results reported here were selected to be elongatingwithin 610% of the mean rate (approximately 3 mm h21

for well-watered and 1 mm h21 for water-stressed roots).

Microtubule Localization

The protocol for immunocytochemical localization of mi-crotubules was described by Liang et al. (1996). Apical20-mm root segments were fixed for 1.5 h in 50 mm Pipesbuffer containing 4% paraformaldehyde, sectioned longi-tudinally at 100-mm thickness on a Vibratome (V-1000,Technical Products International, St. Louis, MO), collectedon slides, and incubated successively in primary (mousemonoclonal against chicken brain b-tubulin, Amersham)and secondary (Cy3-conjugated goat anti-mouse IgG, Jack-son ImmunoResearch Laboratories, West Grove, PA) anti-bodies. For qualitative assessment, sections were peeled offthe slide, leaving cortical cytoplasm affixed to the slide(Liang et al., 1996). Such preparations will be referred to asmicrotubule peels. Quantification was done on sections toavoid the potential for the peeling process to alter micro-tubule angles. To assay the sensitivity of cortical arrays tocold-induced depolymerization, seedlings were put intothin plastic bags (to prevent water uptake) and submergedin ice water (0°C) in a vertical position for 0, 3, 6, or 10 min,and then were fixed and processed as described above.Preliminary experiments showed that cortical arrays werenot visibly affected when roots were placed in plastic bagsand immersed in water at 29°C for 10 min.

To quantify microtubule angles, we selected from eachroot a single longitudinal section near the median. Imagesmade with conventional epifluorescence microscopy werecaptured digitally at 0.5-mm increments along the root,with the boundary between the root cap and quiescentcenter, which was visible in all sections, used as a commonorigin. Microtubule angular distributions were measured

682 Baskin et al. Plant Physiol. Vol. 119, 1999

with an image-analysis program (Image 1, Universal Im-aging, West Chester, PA).

Polarized-Light Microscopy

Apical 6-mm segments were fixed in 50 mm Pipes buffercontaining 1 mm CaCl2 and 4% paraformaldehyde at roomtemperature for 2 h. After the segments were rinsed, theywere embedded in butyl-methyl-methacrylate, as de-scribed by Baskin and Wilson (1997), except that wire loopswere not used because of the large size of the root. Themethacrylate’s refractive index (n 5 1.533, Bayley et al.,1957) matches that of cellulose and other polysaccharidesand thus suppresses “form birefringence,” which resultsfrom the alignment of polymers of one refractive indexwithin a medium of another (Preston, 1974). Sections (2 mmthick) were spread with chloroform vapor and baked onsilane-coated slides for several hours. Measurements of thelength of the root segment before fixation and after embed-ding showed negligible shrinkage, as previously found forthis type of methacrylate (Carlemalm et al., 1982); however,sectioning shortened the length of the root segment byabout 10%, which was not corrected for in the ordinate ofFigure 7.

Sections were mounted in a 75% glycerol, 0.01% TritonX-100 solution and examined through a polarized-lightmicroscope equipped for microphotometry (Jenapol,Zeiss). Longitudinal-radial cell walls that were containedin the plane of approximately median-longitudinal sectionswere examined. The stage was rotated to place the cellwalls at extinction and then rotated by 45°. A suitable cellwall was translated to the optical axis, where a smallsquare aperture (15 mm2 at the magnification used) re-flected 100% of the light to a photomultiplier with a digitalread-out of intensity. Intensities were measured for 645°settings of the compensator (Brace-Kohler type, Gmax 5 18nm). An adjacent background area outside the root wasthen translated to the optical axis, and the intensity wasmeasured at 645° compensator settings. At every 0.5 mmfrom the boundary between the root cap and quiescentcenter, cell walls were measured that were no more than 50mm from the defined position.

Retardation was calculated from the intensity measure-ments with the compensator equations (Jerrard, 1948) asfollows. The intensity through a single birefringent plate at45° between crossed polars (i.e. through the background),Ib, is given by:

Ib 5 Io sin2Sd1

2 D (1)

where Io is the incident light intensity and d1 is the maximalretardation of the compensator (18 nm). The intensitythrough two birefringent plates at 645° between crossedpolars (i.e. through the cell wall), Iw, is given by:

Iw 5 IoSsin2d1

2 cosd2 612sin d1 sin d2 1 sin2

d2

2 D (2)

where d2 is the retardation of the specimen, and the sign ofthe second term is given by the sign of the compensator

(i.e. 645°). The specimen retardation was then obtained bysimultaneous solution of the equations. For convenience,Equation 2 was simplified by using the small angle approx-imation (cosx 5 1; sinx 5 x) for d2. This assumption wasvalidated by finding that the error from the small angleapproximation was less than the precision of the photo-multiplier. The values from the two compensator settingswere averaged to produce a single datum point for eachmeasured cell wall.

Replicas of Cellulose Microfibrils

Apical 20-mm root segments were fixed in 4% parafor-maldehyde in deionized water for 30 min, rinsed in waterseveral times, and sectioned on the Vibratome as describedabove. From each root three approximately median sec-tions were placed in a 20-mL vial and rinsed in distilledwater for at least 2 h. Sections were then incubated in 0.5 mNa2CO3 at room temperature for 3 d to remove pectin, withseveral changes of the solution. After two to three rinses indouble-distilled water, the sections were placed on nitro-cellulose sheets clamped on microscope slides and allowedto dry overnight at 30°C. The preparations were shadowedat an acute angle with platinum, and subsequently fromabove with carbon, in a vacuum evaporator (DV 502, Den-ton, St Louis, MO). Under a stereomicroscope, the shad-owed sections were cut into 0.5-mm segments starting atthe line between root cap and quiescent center. Segmentsoriginating from the same positions along different rootswithin the same experiment and treatment were pooled. Torelease the nitrocellulose and remove tissue remnants, seg-ments were treated with 25% Cr2O3 solution for at least 4 hand then with dilute bleach (about 0.25%) for 1 h. After thereplicas were rinsed extensively, they were mounted on60-mesh hexagonal copper grids and viewed in an electronmicroscope (model 1200, JEOL).

Replicas of longitudinal-radial cell walls of cortical cellswere photographed at 325,000, with the orientation indi-cated by including a transected longitudinal cell wall. Im-ages of the microfibrils were projected through a photo-graphic enlarger onto a circle with a diameter at the level ofthe cell of 600 nm. Each microfibril was traced at thecircumference of the circle, and the angle of the traces withrespect to the root axis was measured using a digitizingtablet and software (SigmaScan, Jandel Scientific, CorteMadera, CA). For measurement, the longitudinal axis wasdefined as 0° and 180°. To average microfibril angles mean-ingfully, the definition of zero was moved in 1° incrementsthrough a total of 180° by addition or subtraction of 180° toappropriate subsets of the data; for each increment, amean 6 sd was calculated and the lowest sd was used toselect the best average, which was then transformed backto its original angle with respect to the longitudinal axis.

To view microtubules and cellulose microfibrils in thesame sections, microtubule peels were prepared from me-dian longitudinal sections as described above, and afterpeeling, the sections were processed for microfibril repli-cas. To ensure that the replicas were made from the sameside of the section from which the microtubule peel hadbeen made, the two sides of the section were identified by

Growth Anisotropy, Microtubules, and Microfibrils 683

cutting the basal end of the section obliquely. Care wastaken to account for all inversions of the image duringelectron microscopy by the use of asymmetric internalfeatures (e.g. numerals on the grid).

For the growth data obtained previously (Liang et al.,1997), the distal end of the root cap was defined as theapex; therefore, to be consistent, for all distances measuredhere with respect to the boundary between root cap andquiescent center, we added the average root cap length(about 500 mm for all treatments), which was measuredfrom approximately median longitudinal Vibratome sec-tions of unfixed roots.

RESULTS

Radial Expansion Rates and the Degree ofGrowth Anisotropy

The first paper in this series (Liang et al., 1997) showedthat the spatial profiles of radial and tangential expansionrates in the maize primary root changed in well-wateredroots with time from germination (“developmental” thin-ning) and also changed in roots transplanted to low waterpotential (21.6 MPa, “water stress”). The developmentalthinning and the thinning induced by water stress resultedfrom decreased radial and tangential expansion rates inboth the cortex and stele. Here we focused on the cortex,although some results were obtained for the stele, becausecortical cells are more homogeneous in shape. Also, for thecortex we considered radial rather than tangential expan-sion rates, because the cell walls expected to limit radialexpansion are longitudinal-radial walls, which can be as-sayed conveniently in median-longitudinal sections. Forthe stele, tangential and radial expansion rates are assumedto be equal (Liang et al., 1997). We did not analyze theepidermis because this tissue was not reliably retained inthe preparations.

To enable the reader to compare readily the spatial pro-files of expansion rate with the results presented below formicrotubule and microfibril orientation, we reproduce rel-evant data from Liang et al. (1997) in Figure 1. The devel-opmental thinning of well-watered roots was analyzed bycomparing them at 24 and 48 h after transplanting. Theprofile of longitudinal strain rate (i.e. relative elementalelongation rate) was similar at these times (Fig. 1A). Incontrast, radial strain rates in the cortex differed: around 3mm from the apex, radial strain rates at 48 h were high,whereas those at 24 h were essentially 0, and between 6 and9 mm from the apex, rates at 48 h decreased to 0, whereasthose at 24 h remained high and even increased (Fig. 1B).Accordingly, the degree of growth anisotropy, calculatedas the ratio of longitudinal to radial strain rate, was strik-ingly different at the two times (Fig. 1C). Anisotropy in-creased with position in the apical 4 mm of the root butreached much higher values at 24 h compared with 48 h;basal of 4 mm, anisotropy decreased steadily at 24 h but at48 h decreased and then increased steeply.

The effect of low water potential was analyzed by com-paring well-watered and water-stressed roots 48 h aftertransplanting, when growth in both treatments had reached

steady state (Liang et al., 1997). Longitudinal strain rateswere similar in the apical 3 mm of the root, and basal of thisthey were reduced in the water-stressed roots (Fig. 1A), aspreviously reported (Sharp et al., 1988). In contrast, radialstrain rates in the cortex were decreased in the apical 4 mmof the water-stressed roots but at more basal locations wereidentical in the two treatments (Fig. 1B). Consequently, thedegree of growth anisotropy in the water-stressed roots wassubstantially higher than in the well-watered roots in theapical 4 mm of the root and basal to this was substantiallylower (Fig. 1C). Thus, both the developmental thinning ofthe roots and the further thinning imposed by water stresschanged radial expansion rates and the degree of growthanisotropy appreciably.

Localization of Microtubules

In median-longitudinal sections, cortical microtubule ar-rays in cortex cells had distinct orientations at defined dis-tances from the apex. Examples are shown of transverse,oblique, and longitudinal orientations for both well-watered

Figure 1. Longitudinal and radial strain rates as a function of dis-tance from the apex for well-watered (ww, 24 and 48 h after trans-planting) and water-stressed (ws, 48 h after transplanting) roots. Thedata are from Liang et al. (1997). A, Longitudinal strain rate. Arrowsindicate positions where microtubule orientation changed fromtransverse to oblique (Fig. 3). B, Radial strain rate for the cortex. C,Growth anisotropy, calculated as the ratio of longitudinal to radialstrain rates. Note that when radial strain rates are near 0 growthanisotropy tends toward infinity; these values are not plotted.


and water-stressed roots (Fig. 2). We reported previouslythat the appearance of obliquely oriented microtubules insections or peels reflects the fact that the cortical arrayforms a helix around the cell; moreover, the handedness ofthe helix at defined positions was conserved among roots(Liang et al., 1996). With increasing distance from the apex,transverse arrays were replaced by right-handed helices,longitudinal arrays, and then by left-handed helices. Thisprogression was the same for well-watered and water-stressed roots, although as shown below, the positionswhere the transitions occurred were different between thetreatments. Comparing the two treatments, microtubulearrays of the same orientation could not be distinguishedvisually.

Quantification of Microtubule Orientation

To determine the extent to which microtubule orienta-tion was associated with the profiles of expansion rate, wequantified the angular distribution of microtubules as afunction of position. Results for the cortex are shown inFigure 3. The net orientation of microtubules was similar inthe 24- and 48-h well-watered roots (Fig. 3A): microtubuleswere transverse until nearly 8 mm from the apex and then

steadily reoriented until they reached a stable orientationof approximately 225° (left-handed helix). For the water-stressed roots, the reorientation of microtubules began andended nearer to the apex. In both treatments the meanorientation of microtubules was transverse throughoutmuch of the growth zone, which is consistent with the factthat elongation rates were consistently greater than radialexpansion rates, i.e. the degree of anisotropy was greaterthan 1 (Fig. 1).

We compared the standard deviation of microtubuleangle to radial strain rates and to the degree of growthanisotropy. In all cases, the deviation among microtubulesslowly increased with distance from the apex until meanmicrotubule orientation was near longitudinal, when thedeviations grew notably (Fig. 3B). Once the mean orienta-tion reached the stable oblique orientation (i.e. 225°), thedeviations again became smaller. Thus, the degree of mi-crotubule alignment tended to change in parallel with thenet orientation of the array but not in relation to thechanged spatial profiles of either radial expansion rate orgrowth anisotropy. For example, between 6 and 9 mm fromthe apex, the deviation in microtubule orientation wassimilar for well-watered roots at 24 and 48 h despite thelarge differences between them in radial expansion ratesand the degree of anisotropy (Fig. 1, B and C).

To extend these results, we compared microtubule ori-entations between well-watered and water-stressed rootsin the stele (Fig. 4). The tangential (radial) strain rates wereconsiderably reduced in the apical 5 mm of the water-stressed stele (Fig. 4A), and the spatial profile of the degreeof growth anisotropy was quite different between the twotreatments (Fig. 4B). In the well-watered roots, anisotropyincreased gradually with position until about 5 mm fromthe apex and then increased steeply to a pronounced max-

Figure 2. Micrographs of cortical microtubules in median-longitudinal sections showing the similar appearance of transverse,oblique, and longitudinal orientations in cells of the cortex of well-watered and water-stressed roots. A to D, Well-watered roots; E to H,water-stressed roots. Examples of transverse (A and E), oblique (B andF; right-handed helical), longitudinal (C and G) , and oblique (D andH; left-handed helical) microtubule orientations. Micrographs wereobtained from peels, as described in “Materials and Methods.” Bar 520 mm.

Figure 3. Microtubule orientation in cortical cells as a function ofdistance from the apex of well-watered (ww) and water-stressed (ws)roots. A, Mean microtubule angle measured for cells localized inmedian-longitudinal sections. B, The SDs of the above distributions.At each position, 20 microtubules were sampled from two cells, andthe data presented were pooled from measurements of 5 to 10 roots(100–200 microtubules measured per position).


imum, whereas in the water-stressed roots anisotropysteadily decreased with position. Microtubules in stelarparenchyma were transverse to even more basal positionsthan in the cortex (Fig. 4C), and as in the cortex the stan-dard deviation of microtubule alignment increased only asmicrotubules became oblique (Fig. 4D). In the first 6 mm ofthe root, the deviations were indistinguishable between thetreatments, despite the large differences in tangential strainrate and in the degree of anisotropy.

To compare clearly the onset of microtubule reorienta-tion with the decrease in longitudinal strain rate, Figure 5re-plots mean microtubule angle against time from themaximal longitudinal strain rate (for the well-wateredtreatment, only the 24-h cortical data are shown for clarity).In the cortical cells of both the well-watered and water-stressed roots, microtubules began to reorient 2 h after thepeak, when the strain rates had already decreased by about25% (Fig. 1A, arrows). In the stele the reorientation waseven later, occurring about 5 h past the peak longitudinalstrain rate for water-stressed roots, when the rate hadnearly reached 0. (The reorientation in the stele was not

defined for the well-watered roots because microtubulepreservation was unreliable beyond about 12 mm from theapex.)

Assessment of Microtubule Stability

Results thus far have shown that the profiles of expan-sion rate were not associated with differences in microtu-bule orientation; however, it is possible that the profileswere associated with differential microtubule stability. Todetermine whether microtubules at different positions or indifferent treatments differed in stability, we fixed rootsafter first exposing them to 0°C for 3, 6, or 10 min. Colddestabilizes microtubules and will lead to their depolymer-ization unless they are stabilized by associated proteins(Bokros et al., 1996). Figure 6 shows that exposure for 6 minsubstantially depolymerized microtubules (compare withFig. 2), but no difference was detected between treatmentsor positions. After a 10-min exposure, very few microtu-bules remained (not shown). Although subtle differences instability would have been missed, these results suggestthat there is not a large population of microtubules withsignificantly altered stability, either as a function of treat-ment or as a function of the type of orientation (i.e. trans-verse, helical, or longitudinal).

Quantification of Cellulose Microfibril Orientation

To quantify the orientation of microfibrils throughoutthe thickness of the wall, we used polarized-light micros-copy to quantify the birefringent retardation of the cellwall. In the cortex the retardation of longitudinal-radial cellwalls of well-watered roots at both times was approxi-mately constant with position (Fig. 7). There was no sug-gestion of increased retardation around 3 mm from theapex in the 24-h treatment that might have accounted forthe greatly lowered radial expansion rate at that location

Figure 5. Mean microtubule angle as a function of time from peaklongitudinal strain rate. Mean microtubule angle for the cortex ofwell-watered (24 h) and water-stressed (48 h) roots and for the steleof the water-stressed roots were re-plotted versus time instead ofposition, taking advantage of the steady-state elongation kinetics andusing the transformation method described by Silk et al. (1984).

Figure 4. Tangential (radial) strain rate, growth anisotropy, and mi-crotubule orientation as a function of distance from the apex for thestele of well-watered (ww) and water-stressed (ws) roots. A, Tangen-tial strain rate. B, Growth anisotropy, calculated as the ratio oflongitudinal to tangential strain rates. Note that the longitudinalstrain rate profile shown in Figure 1A is the same for all tissues. C,Mean microtubule angle measured is stelar parenchyma. D, The SDsof the above distributions. Data in A and B are from Liang et al.(1997) and in C and D are averages of at least 100 microtubulesmeasured per position from 5 to 10 roots.


(and the concomitantly increased growth anisotropy).Polarized-light data were not obtained from more basallocations because adjoining walls may contain microfibrilswith opposite oblique orientations (see below), whichwould complicate interpretation of the data. Comparingwell-watered to water-stressed roots (at 48 h) showed thatwater stress decreased retardation of the cell walls consid-erably (Fig. 7), despite the fact that throughout most of thisregion in water-stressed roots the radial expansion rate wasdecreased and expansion was more anisotropic. In otherwords, the walls of well-watered compared with water-stressed roots expanded faster in the radial direction andless anisotropically, and yet these walls had either morecellulose per unit area of cell wall or more highly alignedcellulose (or both).

Because evidence suggests that only the inner layers ofthe cell wall are load-bearing (Richmond, 1983; Taiz, 1984),we also quantified the alignment of microfibrils in theinnermost layers. Median longitudinal sections were ex-tracted gently to minimize the potential for disruptingmicrofibrils and to ensure that images represented mainlythe most recently deposited layer, and metal-carbon repli-cas were made and examined with electron microscopy.Figure 8 shows a representative image of a cortical cell, inwhich microfibrils are clearly resolved. Because of the dif-ficulty of this method, well-watered roots were analyzed at

only a single time, 24 h. Mean microfibril angle was trans-verse for the first 8 mm from the apex in well-watered rootsand for the first 5 mm in water-stressed roots, as would beexpected if microfibril orientation controlled the directionof maximal expansion rate (Fig. 9A). Basal of these posi-tions, the mean microfibril angle became oblique, and strik-ingly, the sense of the obliquity differed between the treat-ments. As microfibrils became oblique, the direction ofmaximal expansion rate did not change, presumably be-cause cells passed through this region rapidly and stoppedgrowing before synthesizing enough oblique microfibrilsto alter the direction of maximal expansion.

The standard deviation of microfibril angle was corre-lated with neither the observed patterns of radial expansionrate nor the degree of growth anisotropy (Fig. 9B). Forexample, for the well-watered roots, the deviation was vir-tually constant for the first 8 mm despite the differences inradial expansion rate and the degree of anisotropy (Fig. 1,B and C). Results of both methods for examining celluloseorientation concur in showing that the degree of orienta-tion among microfibrils was correlated with neither theamount of radial expansion nor with the degree of growthanisotropy.

Relationship between Orientations ofMicrotubules and Microfibrils

For the diffuse growing cells of higher plants, the orien-tation of microtubules is widely believed to control theorientation of microfibrils. Our results are fully consistentwith this belief where these elements were transverse.However, for the water-stressed roots, comparison of Fig-ures 3 and 9 shows that microtubules reoriented to obliqueangles greater than 90° (right-handed helices) but microfi-brils reoriented to angles less than 90° (left-handed helices).To ensure that this was not a sampling discrepancy be-tween the different experiments, we modified our proce-

Figure 6. Micrographs showing that exposure of the seedlings to 0°Cfor 6 min depolymerized cortical microtubules to the same extent fortransverse, oblique, and longitudinal orientations in well-wateredand water-stressed roots. A to D, Well-watered roots; E to H, water-stressed roots. Examples of transverse (A and E), oblique (B and F;right-handed helical), longitudinal (C and G), and oblique (D and H;left-handed helical) microtubule orientations. Bar 5 20 mm.

Figure 7. Birefringent retardation as a function of distance from theapex of well-watered (ww) and water-stressed (ws) roots. Cortical cellwalls were measured in approximately median-longitudinal sectionsin 2-mm semithin methacrylate sections. Data are means 6 SE of fiveroots, with two cell walls sampled at each position per section andthree sections measured per root.


dures so that microtubules and microfibrils could be exam-ined in the same section. For this analysis, we counted thenumber of cells having orientations judged visually to be inone of the following classes: undefined, transverse, right-handed helical, longitudinal, and left-handed helical. Theorientations of microtubules and microfibrils of water-

stressed roots are compared in Figure 10 by showing thefrequency of cells with a given class of orientation at fourpositions. The orientation of microtubules went from pre-dominantly right-handed helical at 6 mm, through longi-tudinal, to predominantly left-handed helical. However,cellulose microfibrils at 6 and 7 mm from the apex occurredin both helical forms about equally and then became pre-dominantly right-handed helical, which was opposite tothe prevalent orientation of the microtubules.

DISCUSSION

To understand how the degree of anisotropic expansionis controlled, we have studied how the shape of the maizeprimary root changes developmentally and in response towater stress. To our knowledge, this is the only study tohave quantified growth anisotropy in different tissues aswell as orientations of microtubules and microfibrils in thesame material. Our results are consistent with the preva-lent view that the direction of maximal expansion rate isspecified by the mean orientation of microtubules andmicrofibrils; however, we also show that changes in thedegree of growth anisotropy are apparently not specifiedby the orientation of either microtubules or microfibrils.

Figure 8. Electron micrograph showing the appearance of microfibrils on theinnermost layer of a longitudinal-radial cell wall of a cortical cell from awell-watered root. Image shows a cell with a net transverse orientation ofmicrofibrils, approximately 5 mm from the apex. The longitudinal axis of the rootis parallel to the side of the figure. Vibratome sections were extracted withcarbonate and a metal-carbon replica was made as described in “Materials andMethods.” Bar 5 400 nm.

Figure 9. Microfibril orientation as a function of distance from theapex of well-watered (ww) and water-stressed (ws) roots. A, Meanmicrofibril angle measured for cortical cells in longitudinal sections. B,The SDs of the above distributions. Data are averages of 250 to 1800microfibrils measured per position from three experiments with fiveroots each.

Figure 10. In water-stressed roots the handedness of helical orien-tations of microfibrils is in many cells opposite to that of the micro-tubules. Percentage of cortical cells with various classes of orienta-tions of microtubules (MT, white bars) and microfibrils (MF, blackbars) at 6 mm (A), 7 mm (B), 10.5 mm (C), and 11.5 mm (D) from theapex are shown. Percentages of cells with undefined (Un), transverse(Tr), right-handed helical (Rt), longitudinal (Long), and left-handedhelical (Lft) orientations are also shown. Orientations of each ele-ment were measured in the same sections. Data for microtubules aremeans of 100 to 140 cells, and for microfibrils of 270 to 560 cells,from single median-longitudinal sections from five to seven roots.


Orientation and Stability of Cortical Microtubules

Cortical microtubule arrays are well known to changetheir mean orientation in response to various stimuli (Wil-liamson, 1991), but intermediate stages in the reorientationhave rarely been observed. We found that mean microtu-bule orientation changed steadily at an average rate of 1°min21 in well-watered roots and more slowly in water-stressed roots. In pea epidermis, although it took about 45min for the mean microtubule orientation to rotate through45° (i.e. similar to the rate seen here), the reorientation wasdiscontinuous in each cell, with groups of microtubulesreorienting sooner than others (Wymer and Lloyd, 1996). Incontrast, microtubule orientation in both the stele and cor-tex of the maize root remained highly uniform until be-coming longitudinal, when the standard deviation for mi-crotubule angle increased abruptly (Fig. 3B). Our resultsare consistent with both leading models for the mechanismof reorientation (Cyr, 1994; Hepler and Hush, 1996; Wymerand Lloyd, 1996) involving either physical rotation or se-lective stabilization of microtubules.

Our results showing that microtubule arrays of differentorientations and in the different treatments had a uniformsensitivity to cold differ from previous reports. Baluska etal. (1993) reported that the cold stability of microtubules inthe maize root differed as a function of position. For peaepidermis, transversely oriented microtubules were re-ported to be more cold-labile than longitudinal ones(Akashi and Shibaoka, 1987). Also, in the same material,longitudinal orientation and cold stability were both pro-moted by abscisic acid (Sakiyama and Shibaoka, 1990). Thelatter finding is relevant to this study because in the water-stressed maize root abscisic acid accumulates, reaching 5 to10 times the well-watered levels in the apical 5 mm of theroot (Saab et al., 1992). The uniform response of microtu-bules to cold reported here, in contrast to the divergentresponses reported elsewhere, might be explained by theuse of different organs, species, or cultivars; alternatively,the explanation might be the duration of cold treatment,which was 10 min or less here but 1 h or more in the otherstudies. The half-life of cortical microtubules is short, ap-proximately 1 min (Hepler and Hush, 1996); therefore, atreatment that blocks microtubule polymerization shouldremove more than 99% of the cortical array by 10 min.After 1 h or more of cold, cells will have begun to accli-mate, which has been shown to include synthesis of cold-tolerant tubulin isotypes (Chu et al., 1993). Therefore, mi-crotubule arrays present after 1 h or more of cold probablyreflect acclimation rather than microtubule dynamics at theonset of the cold treatment.

Orientation of Cellulose Microfibrils: Polarized-Light andElectron Microscopy

To assess microfibril orientation, we used both polarized-light and electron microscopy of replicas of the innermostsurface of the wall. In recent years studies of the structureof primary cell walls and its relation to growth have fo-cused on the innermost cell wall layer, as revealed byelectron microscopy, because it is now widely believed that

only the innermost layers of the cell wall are load-bearing.This view arose from studies of the giant internodal cells ofcharacean algae, for which evidence was obtained that onlythe inner 25% of cell wall controlled the directional yield-ing behavior of the cell (Richmond, 1983; Taiz, 1984). Since25% of the wall of these extraordinary cells exceeds bymany times the thickness of most primary walls of higherplants, the proportion of load-bearing cell wall thickness inthe internodes may not scale to thinner cell walls. There-fore, the mechanical reinforcement of a primary higherplant cell wall may extend throughout most, if not all, of itsthickness.

For a cell wall, the amount of retardation reflects theamount of crystalline microfibrils in the light path, as wellas their average degree of orientation (Preston, 1974). Re-tardation of cell walls of water-stressed roots was less thanthat of the well-watered roots, whereas microfibril align-ment at the innermost layer of the two treatments hadabout the same variability. This could be explained bymicrofibrils reorienting after deposition as a result of beingrotated passively by the expansion of the cell wall, becausethe amount of this rotation is predicted to be proportionalto the degree of growth anisotropy (Erickson, 1980; Pres-ton, 1982). The greater anisotropy in the apical portion ofthe water-stressed roots would be expected to rotate mi-crofibrils farther away from transverse and, consequently,to decrease retardation, as was observed. However, thepredicted extent of the rotation is modest for the majorityof microfibrils, and it is doubtful that the predicted differ-ence in rotation would amount to a detectable signal. Onthe other hand, the lesser retardation could be explained bycell walls of the water-stressed roots having fewer micro-fibrils per unit area, which would result if water stress haddecreased cellulose synthesis. Regardless of whether thelesser retardation in the water-stressed cortical cell wallsresulted from a decrease in cellulose synthesis or frommore passive rotation of microfibrils, less retardation washypothesized to make growth less anisotropic, which is theopposite of what occurred.

Microtubule-Microfibril Parallelism

In studies of microtubule and microfibril parallelism,most researchers have compared mean orientations andonly a few have compared the variability of alignment. Wefound that microtubules were more highly aligned than themicrofibrils, as judged by the much smaller standard de-viations of their angular distribution (Fig. 3B versus Fig.9B). In contrast, Sassen and Wolters-Arts (1986) reportedthat in stamen hairs of Tradescantia virginiana angular dis-tributions of microtubules were either about the same orlarger than those of the microfibrils (measured in replicas),and Seagull (1992) reported that in cotton hairs the degreeof variability of microtubules was nearly identical to that ofthe microfibrils (in replicas); the variability of both de-creased in parallel from nearly random in young hairs toextremely well aligned in nongrowing hairs. It is possiblethat we underestimated the variability of microtubule ori-entation through imaging microtubules with light micros-copy, because the diameter of a microtubule is 10 times less


than the limit of resolution (Williamson, 1991). Neverthe-less, it is unlikely that this effect could account entirely forthe better alignment of the microtubules. The alignment ofmicrotubules has been hypothesized to affect the alignmentof microfibrils by direct or indirect mechanisms (Emons etal., 1992; Wymer and Lloyd, 1996). For the maize root, ourresults suggest that if microfibril deposition is coupled tomicrotubules directly, e.g. by a motor protein, then thecoupling must be transient to allow for the greater diver-gence among microfibrils.

In addition to the systematic difference in the variabilityof alignments of microtubules and microfibrils, we alsofound a striking difference in their mean orientation. Theorientations of microtubules and microfibrils were parallelwhere these elements were both transverse, but the align-ments diverged when they became helical. This was obvi-ous for the water-stressed roots in which the mean orien-tations of microtubules and microfibrils indicated helices ofopposite handedness, but was also true for the well-watered roots in regions of helical orientation. For exam-ple, in well-watered roots between 12 and 18 mm from theapex, mean microfibril angle was approximately 180°,whereas mean microtubule angle was 225°. Moreover,analysis of different classes of orientation (as reported forwater-stressed roots in Fig. 10) revealed many cells withleft- and right-handed helical microfibril orientation at thesame position (data not shown). In contrast, the orientationof helical microtubule arrays at a given position wasstrictly uniform (Liang et al., 1996). Thus, in both treat-ments, where microtubules were helical, microfibrils werenot always co-aligned. In pea roots mean orientations ofmicrotubules and microfibrils have been found to be par-allel (Hogetsu, 1986; Hogetsu and Oshima, 1986), but it wasnot determined whether oblique orientations reflected he-lices of similar or opposite handedness. Similar to ourfindings, in roots of onion and radish microtubules paral-leled microfibrils when both were transverse, but were notalways parallel when mean orientations were helical (Traasand Derksen, 1989).

Opposite helical alignment contradicts the prevalentview of microtubules aligning microfibrils. A helical align-ment of microfibrils may represent a default orientationstate that can form independently of microtubules (Emons,1994). In some cell types microfibrils are known to bedeposited helically without requiring microtubules, as inroot hairs (Emons et al., 1992) and in some green algae(Mizuta et al., 1989; Kimura and Mizuta, 1994). Most evi-dence supporting a role for microtubules in the alignmentof microfibrils suggests that microtubules are required forcoherent organization of cellulose throughout a cell ortissue but not for the local organization of microfibrils insingle cells or regions of cells. Thus, when microtubules aredepolymerized, there are only a few examples in which thealignment of microfibrils becomes random (Hogetsu andShibaoka, 1978), but many in which microfibrils remainwell aligned locally but lose the consistency of alignmentacross the cell or tissue (Itoh, 1976; Takeda and Shibaoka,1981; Mueller and Brown, 1982). In the maize root it ispossible that the transverse deposition of microfibrils re-quires the presence of transverse microtubules but that, as

microtubule organization becomes helical, microfibril dep-osition becomes uncoupled from microtubules and as-sumes a helical pattern by virtue of some other organizinginfluence.

Growth Anisotropy and the Role of the Epidermis

Our results bear on how microtubules and microfibrilsmay control elongation rates as well as rates of radial andtangential expansion. We found that microtubules and mi-crofibrils reoriented from transverse to oblique after elon-gation rate had already declined significantly, and wefound that the degree of alignment among microtubulesand microfibrils did not explain rates of radial expansion orthe degree of anisotropy. Both of these conclusions mightbe argued against because we did not examine the epider-mis. The epidermis is believed to exert a dominant influ-ence on the elongation of shoots (Kutschera, 1992), and inthe maize root, the outer epidermal wall includes a thickpellicle in the apical 4 to 5 mm of the root and resemblesultrastructurally the outer epidermal cell wall of the shoot(Abeysekera and McCully, 1993a). However, that the epi-dermis limits elongation rates in roots is doubtful. Bjork-man and Cleland (1991) removed the epidermis from maizeroots and found that the spatial profile of longitudinalstrain rate was unaffected.

The epidermis is even less likely to play a role in limitingradial or tangential expansion. Despite the shoot epidermishaving been investigated intensively with respect to elon-gation, to our knowledge, its role in controlling radial andtangential expansion has never been studied. Because dif-ferent tissues expand radially at different rates, as seenhere for the cortex and stele (Figs. 1B and 4A), these ratescannot be limited by a single tissue. Moreover, when thestele expands radially but the cortex does not (as seen herearound 3 mm from the tip), the cortex and the epidermismust nevertheless expand tangentially or be split by theexpanding stele. Thus, at least in the tangential direction,outer tissues need to be compliant to the radial or tangen-tial expansion of inner tissues. In agreement with this, ithas been reported for maize roots that the excised outerepidermal wall is highly compliant tangentially (Abey-sekera and McCully, 1994). Finally, further doubt is cast onthe root epidermis limiting radial or tangential expansionby genetic evidence. In Arabidopsis thaliana (Baskin et al.,1992) and maize (Abeysekera and McCully, 1993b), mu-tants have been characterized in which cells of the rootepidermis are swollen or distorted, and in the maize mu-tant the thick epidermal pellicle is nearly absent; however,except for the distorted epidermis, the roots of the mutantshave the same diameter as those of the wild type.

Microtubules, Microfibrils, and the Control ofGrowth Anisotropy

We have found that the orientations of microtubules andmicrofibrils change predictably during development, fromtransverse in rapidly expanding cells to helical in nongrow-ing cells. A similar developmental sequence for the align-ments of these elements has been reported for various plant


organs, including roots (Hogetsu, 1986; Hogetsu and Os-hima, 1986). However, from previous reports one could notdetermine whether the reorientation of microtubules andmicrofibrils preceded or followed the decrease in elonga-tion rate because the spatial profile of elongation rate wasnot measured with sufficient accuracy. In contrast, Prit-chard et al. (1993) measured elongation of the maize rootwith enough spatial accuracy but measured microfibrilangles at only four locations. Our results show that as cellsmoved through the growth zone, the reorientation of mi-crotubules and microfibrils followed the decrease in elon-gation rate by 2 h in the cortex and by even longer in thestele. Evidently, the developmental changes in the orienta-tions of microtubules and microfibrils do not cause thechanges in the rate of elongation.

Concerning radial and tangential expansion, we under-took this investigation to learn how the degree of expan-sion anisotropy is controlled. We hypothesized that thiscontrol is exerted by the degree of alignment among cellu-lose microfibrils. This is an economical hypothesis in thatmicrofibrils would control the direction of maximal expan-sion through their mean orientation, as well as the degreeof anisotropy through the dispersion around the meanorientation. This hypothesis was first made by Green(1964), who compared two algae, Nitella axillaris and Hy-drodictyon africanum, and found that the greater degree ofgrowth anisotropy in N. axillaris was associated with morehighly aligned microfibrils. Although Green’s data mayindicate that the hypothesis is sometimes true, instead theymay reflect fortuitous differences in wall structure betweenthe two divergent algal species. Also consistent with thehypothesis, Probine (1965) found that as the diameter ofexcised pea epicotyls increased in the presence of increas-ing concentrations of cytokinin, regions of the cell wallwith transverse microfibrillar orientation had decreasedretardation. However, the increased diameter may havebeen caused instead by bands of longitudinal microfibrilsthat appeared in hormone-treated material and increasedin prominence with concentration. For the maize root,we falsified the hypothesis by finding that changes inthe degree of growth anisotropy were accompanied by aconstant degree of alignment among microfibrils, boththroughout the cell wall and at the innermost layer.

If the degree of expansion anisotropy is not controlled bythe degree of alignment among microfibrils, then whatdoes exert this control? We hypothesize that cell wall yield-ing is regulated independently in longitudinal and radialdirections. Such independent regulation would occur iflongitudinal extensibility were regulated by specific cellwall components that resist the separation of microfibrils,whereas radial extensibility would be regulated by othercomponents that resist shear between microfibrils. Consid-erable evidence suggests that elongation is limited at leastto some extent by the network of hemicellulose that en-meshes microfibrils (Cosgrove, 1997). However, as Liang etal. (1997) pointed out, for the treatments studied here, theactivities of two enzymes, expansin and xyloglucan endo-transglycosylase, thought to loosen this network, are cor-related with longitudinal strain rates but not with radial ortangential strain rates. Although cell wall components that

are active radially have not been identified biochemically,they may have been identified genetically. Tsuge et al.(1996) identified two loci in A. thaliana that exert indepen-dent control of expansion in length and width of leaves.Two other A. thaliana loci have been identified that arerequired for highly anisotropic growth in roots not for thetransverse orientation of microtubules or microfibrils (A.Wiedemeier, T.I. Baskin, unpublished data). Possibly, suchloci encode activities that regulate the ability of alignedmicrofibrils to resist shear stress.

Our results show that in higher plants, unlike in the N.axillaris internode, the degree of growth anisotropy varies,and this variation produces adaptive changes in organshape, such as the thinning of roots in response to waterstress. The extent of growth anisotropy has been neglectedby models relating cell wall architecture to expansion,which have dealt solely with elongation (Cosgrove, 1997).These models must be extended to encompass the fullthree-dimensional yielding behavior of the cell wall beforeplant morphogenesis can be fully understood.

ACKNOWLEDGMENTS

We thank Jan Wilson for flawless technical assistance andCorine van der Weele for thoughtful comments on the manuscript.

Received July 22, 1998; accepted November 7, 1998.

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regulation of growth anisotropy in well-watered and water

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